Control of food spoilage fungi by ethanol

Food Control 22 (2011) 360e368

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Control of food spoilage fungi by ethanol Thien Dao, Philippe Dantigny* Laboratoire de Génie des Procédés Microbiologiques et Alimentaires, Université de Bourgogne, Agro-Sup Dijon, France

a r t i c l e i n f o

a b s t r a c t

Article history: Received 11 March 2010 Received in revised form 2 August 2010 Accepted 10 September 2010

This review discusses the effects of ethanol on the inhibition of growth and germination of fungi and on the inactivation of fungal spores. After a brief survey on the impact of spoilage fungi on the economy and food quality, the major applications of ethanol in controlling fruit decay and extending the shelf-life of food products are reviewed. Many parameters including minimum inhibitory concentration (MIC) and D-values for various moulds are included. The thermodynamic relationship between the liquid phase and the headspace and the mode of action of ethanol on fungi are explained. Due to their promising use as a fumigant, special attention is paid to ethanol vapours. Ó 2010 Elsevier Ltd. All rights reserved.

Keywords: Ethanol Conidia Fungi Mould Inhibition Inactivation Predictive mycology

1. Introduction Moulds can grow on a great variety of substrates and a wide range of pH, water activity (aw) and temperature. Fruits juices are usually quite acid, in the range pH 1.8e2.2 (lemons) to 4.5e5.0 (tomatoes), but the rind tissue that fungal postharvest pathogens colonize is about pH 5 to 5.5 (lemons). Tissue pH alone does not account for the dominance of fungi as plant pathogens. Fungi are capable of rapid invasive growth into tissue, they have developed environmental sensing mechanisms, enabling them to tailor in ambient conditions, by acidification and alkalinisation, to best fit their offensive arsenal (Prusky & Yakoby, 2003). Fungi are capable of rapid invasive growth into tissue while bacteria are not. Therefore microbial spoilage of fruit and fruit products is always caused by fungi (Pitt & Hocking, 1999). Most Penicillia can develop as low as pH 2 (Panasenko, 1967) and many are postharvest pathogens. Almost all xerophilic fungi (i.e., capable of growth at 0.75 aw) are ascomycetes that produce ascospores in asci. Amongst the ascomycetes, Eurotium species are the most common causes of spoilage of dried cereals. Other ascomycetes (e.g., Byssochlamys species, Eupenicillium lapidosum, Neosartorya fischeri, Talaromyces species) are producing ascospores of very high heat resistance which can * Corresponding author. Laboratoire de Génie des Procédés Microbiologiques et Alimentaires, AgroSup Dijon, 1 Esplanade Erasme, 21000 Dijon, France. Tel.: þ33 (0) 3 80 70 44 71. E-mail address: [email protected] (P. Dantigny). 0956-7135/$ e see front matter Ó 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.foodcont.2010.09.019

survive heat processing and are responsible for spoilage of pasteurized foods. In addition, most of the moulds with the notable exception of the Aspergilli exhibited a minimum temperature for growth close to the freezing point. Moulds can develop at the field level. It has been reported that 25% of agriculture products are contaminated with mycotoxins (Mannon & Jonhson, 1985). But moulds can also develop during storage of raw products, and subsequent transport and sale causing considerable economic losses annually for food manufacturers and consumers alike. It is very difficult to assess losses attributable to moulds. In the baking industry, these losses varied between 1% and 3% of products depending on season, type of product and method of processing (Malkki & Rauha, 1978). Another estimate from one bakery in the US was 5% losses (Killian & Krueger, 1983). Even assuming only 1% losses, moulds could be spoiling over 23,000 tons of bread worth nearly £20 million in the UK every year. Throughout Western Europe the annual losses could be around 225,000 tons of bread worth £242 million (Legan, 1993). More generally losses of food to fungal spoilage in Australia must be in excess of $10,000,000 per annum: losses in damp tropical climate and countries with less developed technology remain staggering (Pitt & Hocking, 1999). In the fruit industry, postharvest losses are 5e10% when postharvest fungicides are used (Cappellini & Ceponis, 1984). Without fungicides, losses of 50% or higher have occurred in some years. For example, in a 1993 test to assess the decay potential of stone fruit, an average of 52.8% (range 15e100%) of the fruit decayed during the ripening of eight collections that had not been

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treated with postharvest fungicides (Margosan, Smilanick, Simmons, & Henson, 1997). Moulds are disseminated through spores that are produced in the environment. Air can be a vector in the distribution of almost all fungi that are relevant to food spoilage but packaging materials may also be source of fungal contamination (Scholte, 1995). It is troublesome that after germination of the spores, moulds spread rapidly by aerial mycelia along the fruits, cereals, and food. In order to delay mould growth and subsequent production of metabolites, amongst which the most important for food quality and safety are mycotoxins (D’Mello & MacDonald, 1997) it would be highly desirable to control food spoilage fungi. Among the physicochemical hurdles cited in the literature, ethanol is recognized as a mould inhibitor (Legan, 1993). For a long time, ethanol has been used as a fungicide treatment. Ethanol is used commercially in a lot of products, such as perfumes, many food products paints, alcoholic beverages and additives. Ethanol is a small molecule produced either by chemical synthesis or by microbial fermentation. Recently, there has been substantial interest in nonbiological control agents as well as biological control agents to replace the existing chemical applications. Non-biological control involves chemicals that are Generally Regarded as Safe “GRAS” product such as ethanol (Karabulut, Mlikota Gabler, Mansour, & Smilanick, 2004; Romanazzi, Karabulut, & Smilanick, 2007) as alternative treatments. As such it can be used in the food industry. Ethanol is also easily miscible with water in all proportions. It is a good solvent, as starting compound for the manufacture of dyes, cosmetic and explosives. Ethanol is a familiar constituent of many beverages and is considered to be the least toxic of the straight-chain alcohols (methanol, ethanol, propanol, butanol, etc) evaporates quickly at room temperature, not leaving any residues (Nittérus, 2000). For a long time, it is also as a disinfectant due to its antibacterial properties. Ethanol is a common food component with potent antimicrobial activity (Feliciano, Feliciano, Vendruscuolo, Adaskaveg, & Ogawa, 1992; Karabulut, Smilanick, Mlikota Gabler, Mansour, & Droby, 2003; Larson & Morton, 1991). Smith (1947) concluded that 95% ethanol was best for wet surfaces, 50% for dry surfaces, and 70% for either wet or dry surfaces. This percentage has been shown to be a superior surface disinfectant properties to control bacteria in hand washing, skin de-germing, or instrument sanitation tests (Ali, Dolan, Fendler, & Larson, 2001). But there is no evidence that this ethanol concentration is also optimum to kill fungi. The inhibitory effect can be obtained by adding ethanol directly to the product or by using ethanol vapours. Many studies were concerned with the use of ethanol for extending the shelf-life of food products and for reducing fruit decay. In a first section, these studies will be reviewed. Although this literature did provide useful guidelines for preserving food and agriculture products most of the previous studies were concerned with natural microflora, including bacteria, yeasts and moulds. Depending on its concentration, on the type of product and organisms, ethanol may have different effects on mould development. In a second section, the inhibitory effects of ethanol on growth and germination and on the inactivation of spores will be reviewed. In contrast to the previous section, these studies were concerned with some moulds isolated from spoiled products and cultivated in pure cultures. The effects of ethanol (either through liquid application or vapours in the headspace) will be assessed under controlled conditions in terms of ethanol concentration, temperature, pH and water activity. In fact, it was pointed out that when ethanol vapour generators were used for preserving foods, the concentration of ethanol in the headspace was not constant (Daifas, Smith, Tarte, Blanchfield, & Austin, 2000; Daifas et al., 2003). In the third section the application of liquid ethanol will be compared to the use of ethanol vapours in terms of origin of the

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contaminations, regulations and efficacy of the treatments. There is much controversy about the mode of action of ethanol. Seiler (1989) found no difference in the inhibitory effect of ethanol applied as liquid or as vapour. In contrast, Smith et al. (1987) reported that lower levels of ethanol were required in the vapour phase compared to direct addition to a medium or food for complete microbial inhibition. According to Lerici and Manzocco (2000), the toxicity of ethanol can be described by its ethanol vapour pressure. This assumption is in accordance with the increase of ethanol toxicity with temperature. However, recent studies (Dantigny, Dao, Dejardin, & Bensoussan, 2007) have shown that the toxicity of ethanol cannot be explained by its vapour pressure only. The mode of action of ethanol should also be evaluated at the cell level. The fourth section aimed at reviewing the biological principles for the effects of ethanol on fungi.

2. Industrial use of ethanol 2.1. Extension of mould free shelf-life Addition of ethanol increases the mould free shelf-life of bread when added after baking and cooling at concentrations from 0.5 to 3.5% (wt/wt) of loaf weight (Geiges & Kuchen, 1981; Plemons, Staff & Cameron, 1976; Seiler, 1984; Seiler & Russell, 1991; Vora & Sidhu, 1987). Differences in methods between workers make it difficult to compare results. The data of greatest practical use are those of Seiler (1984), which show that the shelf-life increases with ethanol concentration, a 50% extension in life being obtained with the addition of 0.5% ethanol based on loaf weight. Ethanol is only permitted at levels up to 2% by product weight in pizza whether the alcohol is added directly to the food or not. In Italy, sandwich loaves named “pancarrè per tramezzini” contain up to 2% ethanol wt/dry wt and do not need any addition of sorbic or propionic acid (Bonetto & Bortoli, 1996). Food products such as cake and bread treated with >2% ethanol (wt/wt) were rejected by consumers on the basis of flavour and/or odour (Seiler, 1978). A level of 2% ethanol in food can extend the mould free shelf-life of products but is generally not sufficient to prevent mould development. The inhibitory effect of ethanol can also be obtained by using an encapsulated ethanol pouch (Ethicap, Freund Industrial Co. Ltd., Tokyo, Japan). This ethanol emitter extended the shelf-life of packaged apple turnovers (Smith et al., 1987), pita bread (Black, Quail, Reyes, Kuzyk, & Ruddick, 1993), packed sliced rye bread (Salminen et al., 1996), English-style crumpets (Daifas et al., 2003) and pre-baked buns (Franke, Wijma, & Bouma, 2002). The effects of ethanol on a complex mycoflora were difficult to assess because the ethanol in the headspace was not constant throughout the experiments. This was due to the absorption of ethanol by the products. Therefore the effect of ethanol could also be attributed to the ethanol absorbed by the product. It was shown that the concentration of ethanol required for inhibition was dependent on the water activity of the system (Smith et al., 1987). In fact, it was shown that both aw and moisture affected the vapour pressure of ethanol as a consequence of water-solute and ethanolesolute interactions in the matrix. These interactions varied according to the modality of equilibration (desorption or absorption) at a given aw (Pittia et al., 2006). In addition, the use of ethanol emitter was related to modified atmosphere packaging and it was suggested that ethanol vapours could also affect the film’s permeability to oxygen (Black et al., 1993). The empirical studies that have used ethanol for either increasing the mould-free shelf-life of food products or controlling postharvest decay of fruits are reported in Table 1.

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Table 1 List of empirical studies that have used ethanol for either increasing the mould free shelf-life of food products or controlling postharvest fruit decay. Product

Treatment

Concentrations

Citation

Table grapes Table grapes Peaches Pre-baked buns

Vapour Vapour Vapour Ethanol emitter Incorporation Immersion Immersion Immersion Immersion Immersion Vapour

5 ml/kg grape 2 ml/kg grape 20e45% 0.55e2.2 g/140 g bun 2e12% (v/v) 35, 50% (v/v) 20, 50% 10, 20% 20e70% 20e50% 20e70%

Chervin et al., 2003 Chervin et al., 2005 El-Sheik Aly et al., 2000 Franke et al., 2002 Geiges & Kuchen, 1981 Karabulut et al., 2003 Karabulut et al., 2004 Karabulut et al., 2005 Lichter et al., 2002 Lichter et al., 2005 Lihandra, 2007

Vapour Immersion Immersion

4, 8 ml/kg grape 5e20% 10, 20%

Lurie et al., 2006 Margosan et al., 1994 Margosan et al., 1997

35% 0.5e3.5% 10, 20% 0.33e1.65 g/215 g bread 0.2e1.4% 0.5e3.5% 2.5e40% 2.2 g/100 g apple

Mlikota Gabler et al., 2005 Plemons et al., 1976 Romanazzi et al., 2007 Salminen et al., 1996 Seiler, 1984 Seiler & Russell, 1991 Smilanick et al., 1995 Smith et al., 1987

10, 20% (w/v) 0.5e2% 0.06e0.16% (v/v) 200e1500 mL/L

Spadaro et al., 2004 Vora & Sidhu, 1987 Yuen et al., 1995 Zhang et al., 2007

Bread Grapes Table grapes Table grapes Table grapes Grapes Oranges and peaches Table grapes Strawberries Peaches and nectarines Table grapes Bread Table grapes Sliced rye bread

Immersion Incorporation Immersion Ethanol emitter Bread Incorporation Bread Incorporation Lemons Immersion Packaged apple Ethanol turnovers emitter Apples Immersion Bread Incorporation Oranges Vapour Chinese Vapour bayberries

solutions 5e10% at 22  C for 1e4 min had no effect on the decay due to the prevalent Rhizopus stolonifer, the decay was reduced from 77% (control) to only 1% after treatment with 10% ethanol for 4 min at 45  C, Table 2. But, the same treatment did reduce the decay to only 36% for “Swede” strawberries infested by the prevalent Botrytis cinerea (Margosan et al., 1994). These results clearly indicate that the sensitivity to ethanol is greatly dependent on the mould. In a more recent study, peaches dipped into 20%e100% ethanol solutions were completely rotten by ten days when stored at room temperature, but the peaches experienced little to no browning. In contrast, untreated and fungicide-treated fruit were protected for 1d and 2d, respectively (Lihandra, 2007). The effects of ethanol vapour on the growth of Penicillium italicum and Penicillium digitatum on oranges were assessed by Yuen, Paton, Hanawati, and Shen (1995). These authors reported that exposure to 0.16% ethanol vapour for 5d delayed the appearance of infection symptoms caused by P. digitatum and P. italicum by about 10d and 8d, respectively. The use of ethanol in food at low concentrations (approximately 2% wt/wt) may inhibit growth and germination thus extending the free shelf-life of food products. In contrast, at higher ethanol concentrations, it is suggested that a significant fraction of the spores present at the surface of the dipped fruits were inactivated. However the exposure to ethanol did not appear sufficient for a complete inactivation because fruits were eventually rotten. The more recent studies aimed at optimizing the combination of ethanol concentration, temperature and duration of the treatment without altering the quality of fruits. 3. Liquid treatments 3.1. Effect of ethanol on fungal growth

2.2. Control of postharvest decay of fruits Ethanol dips and vapours were reported to control postharvest diseases of apple (Spadaro, Garibaldi, & Gullino, 2004), peaches (ElSheik Aly, Baraka, & El-Sayed Abbass, 2000; Feliciano et al., 1992; Margosan et al., 1997), lemons (Smilanick, Sorenson, & Henson, 1995), Chinese bayberries (Zhang et al., 2007) and table grapes (Chervin et al., 2003; Chervin, Westercamp, & Monteils, 2005; Karabulut et al., 2003; Karabulut, Romanazzi, Smilanick, & Lichter, 2005; Lichter et al., 2002; Lichter, Zutahy, Kaplunov, Aharoni, & Lurie, 2005; Lurie et al., 2006; Mlikota Gabler & Smilanick, 2001; Mlikota Gabler, Smilanick, Aiyabei, & Mansour, 2002; Mlikota Gabler, Smilanick, Ghosoph, & Margosan, 2005; Romanazzi et al., 2007) especially when heated (Margosan, Smilanick, & Simmons, 1994; Margosan et al., 1997; Smilanick et al., 1995). Whereas immersion of naturally-infested “Chandler” strawberries in ethanol

Geiges and Kuchen (1981) have assessed the influence of ethanol on mycelium development for some moulds grown in pure cultures. Experiments were carried out by adding ethanol to bread slices inoculated with spores. Growth inhibition was observed qualitatively for ethanol concentrations in the range 0e12% (v/v) depending on the mould. In the range 0e4%, growth was observed for all the moulds studied (i.e., Penicillium expansum, Penicillium implicatum, Penicillium lanoso-coeruleum (¼ Penicillium aurantiogriseum), Aspergillus oryzae and Trichoderma harzianum). But growth was arrested at 8% for all these moulds. In contrast, Perry and Beale (1920) found that Penicillium glaucum (¼ most probably P. expansum) would grow in concentrations up to 8% alcohol in dextrose broth. Another study reported that the inhibitory concentration of ethyl alcohol for Penicillium carmine-violaceum (¼ P. roseopurpureum), P. citrinum and P. glaucum (¼ most probably P. expansum) were in the range 5e5.5%

Table 2 Effect of hot ethanol treatments on postharvest control of gray mould (Botrytis cinerea) and black rot (Rhizopus stonolifer) of strawberries. Selected data from Margosan et al. (1994). Strawberry (Control test)

Prevalent pathogen

Test

T ( C)

Decay (%) Immersion time (min)

Chandler (Untreated Dry Control: 77% decay)

Rhizopus stolonifer

Swede (Untreated Dry Control:97% decay)

Botrytis cinerea

Water 10% ethanol Water 10% ethanol Water 10% ethanol Water 10% ethanol

22 22 45 45 22 22 45 45

1

4

77 72 55 29 99 100 97 96

78 80 10 1 100 99 62 36

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(Krause & Ellis, 1937). Biomass production by Pyrenophora avenae was reduced by 20% in 1% ethanol, growth in 3% and 6% ethanol reduced biomass by 57 and 95% respectively (Walters, McPherson, & Cowley, 1998). A non-linear model was developed for assessing the effect of ethanol on the growth of twelve different moulds on Potato Dextrose Agar (0.99 aw) at 25  C (Dantigny, Guilmart, Radoi, Bensoussan, & Zwietering, 2005). This model was able to estimate the minimum inhibitory concentrations, MIC for growth and the concentrations of ethanol at which the radial growth rate, m was KðMICEÞ ; where mopt was the equal to mopt/2, K : m ¼ mopt K:MIC2K:EþMIC:E growth rate at 0% ethanol. The model fit the experimental data with a good accuracy. It was capable of describing curves, m vs. E, with either a concave shape (K < MIC/2) or a convex one (K > MIC/2). MIC was estimated in the range 3%e5% for all moulds with the notable exception of T. harzianum (MIC ¼ 2.14%) and Paecilomyces variotii (MIC ¼ 6.43%), Table 3. These results demonstrated that except T. harzianum, no moulds were completely inhibited by ethanol concentrations of 2% in foods. Aspergillus niger and Mucor racemosus were characterised by equivalent minimum inhibitory concentrations of about 4.2%. But the K values were fairly different, 3.82% and 1.57% for A. niger and M. racemosus, respectively. While 2% ethanol would decrease the growth rate of M. racemosus to about a third of mopt, this concentration would have almost no inhibition effect on A. niger.

3.2. Effect of ethanol on germination In studies assessing the effect of ethanol on germination, spores are exposed to ethanol throughout the experiments. These studies aimed at determining the minimum inhibitory concentration for germination. The underlying concept is that prevention of germination would also prevent from growth. Because many studies assessing the influence of ethanol on fungal growth were carried out using spores as the inoculum, it can be assumed that no growth was observed because spores were unable to germinate. Accordingly, no difference was shown between the minimum inhibitory concentrations for growth and for germination for some Penicillia (Krause & Ellis, 1937). A complete inhibition of germination of conidia from these Penicillia was achieved at 50 g l1. However, it would be possible that some spores (conidia) were germinated but eventually were no longer capable of producing hyphae. While the minimum ethanol concentration for growth of Penicillium chrysogenum was estimated to 3.93% (Dantigny,Guilmart et al. 2005), the minimum ethanol concentration for germination was estimated to 4.3% (Dantigny, Tchobanov, Bensoussan, & Zwietering, 2005). In the latter study, some spores were producing a germ tube

Table 3 Minimum inhibitory concentrations (MIC) and ethanol concentrations at which the growth rate is halved, (K) for various food spoilage fungi. Adapted from Dantigny, Guilmart, et al. (2005) Mould

K (%, wt/wt)

MIC (%, wt/wt)

Aspergillus candidus Aspergillus flavus Aspergillus niger Cladosporium cladosporioides Eurotium herborarium Mucor circinelloides Mucor racemosus Paecilomyces variotii Penicillium chrysogenum Penicillium digitatum Rhizopus oryzae Trichoderma harzanium

2.13 1.57 3.82 1.86 1.71 1.43 1.33 3.25 2.22 0.71 1.63 1.16

3.92 4.60 4.22 3.58 3.25 4.09 4.13 6.43 3.93 3.36 4.79 2.14

363

(synonymous for germination) that eventually blown up. More interestingly, after 3 weeks, the cultures that failed to germinate were allowed to continue to incubate. The ethanol solutions of 4e6% (wt/wt) were substituted for a 9 g/L NaCl solution, and the cultures were incubated at 25  C. In all cases, visible mycelium appeared during the next days, indicating that many conidia remained viable. Under these experimental conditions, the inhibitory effect was clearly reversible. 3.3. Influence of ethanol on inactivation Micro-organisms should be killed rather than inhibited, so it is important to know which ethanol concentrations will result in inactivation. It was shown that exposure to 40% ethanol completely inhibited the germination of B. cinerea (Lichter et al., 2002), but the immersion time was not stated. In another study, it was reported that no germination of B. cinerea was observed on potato dextrose agar (PDA) after immersion in 30% ethanol for 10s at 22e24  C (Karabulut et al., 2004). Germination of B. cinerea spores on PDA after a 30s immersion in 10 or 20% ethanol was 87 and 56%, respectively, compared to 99% among untreated controls (Karabulut et al., 2005). The incidence of grey mould (caused by B. cinerea) infected grape berries that were untreated, or immersed for 1 min in ethanol 35% at 25 or 50  C, was 78.7, 26.2, and 3.4 berries/kg, respectively, after 1 month of storage at 0.5  C and 2 days at 25  C (Mlikota Gabler et al., 2005). Fifty percent ethanol can inactivate Sclerotina fructicola conidia in 0.08 min on peach fruit surfaces and 60% ethanol killed Gilbertella persicaria and R. stolonifer sporangiospores on uninjured surfaces of peach fruits in 1min (Ogawa & Lyda, 1960). But the same authors reported that, on injured surfaces, 70% ethanol for 40 min was needed to kill Gilbertella and 70% for 60 min to kill Rhizopus. Margosan et al. (1994) demonstrated a synergistic effect of heat and ethanol when they reported that diluted ethanol treatments controlled postharvest decay of strawberries caused by B. cinerea and R. stolonifer. A similar approach, employing exposure to 10e20% (v/v) ethanol at 46e50  C for up to 2.5 min, controlled green mould on lemons, caused by P. digitatum and brown rot caused by Monilinia fructicola on peaches and nectarines (Margosan et al., 1997; Smilanick et al., 1995). Temperature has marked influence on ethanol toxicity and its effectiveness to control postharvest decay. Heat and ethanol combinations were synergistic in the control of some fungi (Dao, Bensoussan, Gervais, & Dantigny, 2008) and the diseases they cause facilitating the use of lower temperatures and reduced ethanol concentrations when these approaches are combined. The influence of temperature during a 30 s exposure on the in vitro toxicity of ethanol, expressed the concentrations (%, vol/vol) that inhibited the germination of the treated population by 50% (EC 50) was reported by Mlikota Gabler, Mansour, Smilanick, and Mackey (2004). At 25  C, EC50 concentrations to R. stolonifer, A. niger, B. cinerea and Alternaria alternata were 40, 36, 25, and 20% respectively, while at 40  C, the EC50 concentrations were 20, 20, 14, and 10% respectively. Ethanol toxicity to conidia of these fungi increased about two-fold between 25 and 40  C. By applying these treatments, injury to the fruits did not occur, no off-flavours or odours were detected by the investigators, and an increased firmness of the fruits was observed in most tests (Margosan et al., 1997). These authors also reported that peaches dipped in 20%e100% ethanol experienced little to no injury to the fruit. Ethanol vapours improved apple appearance because they were shown to reduce scald on apples (Scott, Yuen, & Ghahramani, 1995). A complete inhibition of germination does not mean necessarily that all spores are inactivated. Non-thermal inactivation, such as

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that caused by ethanol, of a homogeneous population of microorganisms can be described with a D-value. The D-value, the decimal reduction time, is the time that is needed to inactivate 90% of the micro-organisms. From the data of Geiges and Kuchen (1981), the D-values were calculated and reported in Table 4 for different percentage of ethanol. In contrast to the other studied mould, P. lanoso-coeruleum (¼ P. aurantiogriseum) was insensitive to 10% ethanol. Ethanol was effective from 13% and beyond. T. harzianum was the less resistant mould to ethanol in accordance with previous results on the influence of ethanol on mould growth (Dantigny,Guilmart et al. 2005). At 19% ethanol, P. implicatum was the most resistant to ethanol, thus suggesting a greater z-value. The z-value (% ethanol) can be considered as the increase in ethanol concentration achieving the same inactivation within 10% of the previous exposure time.

The water activity due to ethanol, awEtOH, and the water activity due to glycerol, awGly, can be derived from the Langmuir equations (Lerici, Nicoli, & Manzocco, 1996):

mH2 O 0:438awEtOH ¼ mEtOH 1  0:99awEtOH

(4)

0:236awGly mH2 O ¼ mGly 1  0:99awGly

(5)

The water activity of the solution is given by the Ross equation:

awSol ¼ awEtOH $awGly

(6)

The relative humidity RH (%) in the headspace depends on the water activity of the solution according to:

4. Vapour treatments

RH ¼ 100awSol

4.1. Thermodynamic relationship

Solving the equations, the composition of ternary solutions can be calculated for setting the ethanol vapour pressure and the relative humidity at the desired values physics permitted.

In a hermetically closed vessel, there are relationship between the headspace and the liquid solution at steady state. Therefore it is not necessary to measure the vapour pressure of ethanol in the headspace. The liquid may be a binary water/ethanol or a ternary water/ethanol/glycerol solution. In addition to modifying the ethanol vapour pressure, the addition of glycerol allows the control of the relative humidity in the headspace. To maintain the same humidity in the headspace at different ethanol concentrations in the liquid it is necessary to adjust the aw of the solution by adding glycerol (or other aw depressant). For a ternary solution that contains water (g), mH20; ethanol (g), mEtOH; and glycerol (g), mGly, the molar fraction of ethanol in the liquid is :

Xe ¼

mEtOH 46

(1)

mEtOH mGly mH2 O þ þ 46 92 18

The ethanol vapour pressure, Pe (kPa), in the headspace is:

Pe ¼ Xe ge Pt

(7)

4.2. Effect of ethanol on fungal growth Exposing fruit to ethanol vapours proved effective at inhibiting fungal growth. Peaches that were exposed to 70% or 100% ethanol vapours were protected against fungal infection for up to 30 days when stored at either 4  C or room temperature. In contrast, at room temperature, untreated peaches and fungicide-treated peaches were spoiled by fungi after 2 and 3 days, respectively. 20% ethanol protected peaches for ten days when stored at 4  C and two days at room temperature. Oranges that were exposed to 20%, 50%, 70% and 100% ethanol vapours were protected from fungal infection for 30 days at both 4  C and room temperature (Lihandra, 2007). The use of ethanol vapours at a dose rate of 2 ml kg 1 to limit the development of B. cinerea on table grapes was also reported (Chervin et al., 2005). The total inhibition of the growth of Penicillium notatum, corresponding to the MIC of ethanol in vapour phase, was observed at a concentration of 8.6 mmol/L air (Tunc, Chollet, Chalier, Preziosi-Belloy, & Gontard, 2007).

(2)

where ge the activity coefficient that can be considered equal to 1 for Xe less than 0.8 (Clausen & Arlt, 2002), Pt (kPa) the ethanol vapour pressure at saturation. Pt depended on T ( K) according to a Claudius-Clapeyron law that was approximated by plotting Ln (Pt) vs. (1/T), [data from Lide (1995)]:

Ln ðPtÞ ¼ 19:11  5094=T

(3)

 C,

At 25 (298 K) Pt ¼ 7.5 kPa. Assuming the atmospheric pressure 105 Pa, 1 kPa ethanol vapour pressure is equivalent to 1% (v/v) ethanol in the headspace.

Table 4 Influence of ethanol on the inactivation of conidia of different fungi. D-values were calculated from the data of Geiges & Kuchen, 1981. Mould

D6% (d) D8% (d) D10% (d) D13% (d) D16% (d) D19% (d)

Penicillium expansum Penicillium implicatum Penicillium aurantiogriseum Aspergillus oryzae Trichoderma harzianum

ni 34.7 ni ni ni

ni 15.3 ni 87.2 16

11.8 11.1 ni 19.2 3.3

3.1 9.7 36.1 4.7 2.8

ni: less than 1log reduction in viable conidia after 100d.

1.7 7.8 13.1 2.7 0.9

0.8 3.9 1.4 1.4 0.5

4.3. Influence of ethanol on germination The first detailed study of the inhibitory effects of ethanol vapour on the germination of P. glaucum (¼ most probably P. expansum) and Sterigmatocystis nigra was early demonstrated by Lesage (1895, 1897). The inhibition of the germination of P. chrysogenum conidia by ethanol vapours was studied in the range 0e4% (wt/wt) (Dantigny, Tchobanov et al. 2005). It was shown that all conidia of P. chrysogenum germinated at ethanol concentrations that were 3.0% (wt/wt) or less. At 3.5% (wt/wt), about 60% of the inoculated spores were capable of germinating after 40 h, eventually forming a mycelium. After 3 weeks of incubation, no germination had occurred at ethanol concentrations of 4% and higher. Ethanol vapour was also effective in retarding the apparition of green mould and blue mould on oranges (Yuen, Paton, Hanawati & Shen, 1995). The authors reported that exposure to 0.16% ethanol vapour for 5d delayed the appearance of symptoms caused by P. digitatum and P. italicum, by 10d and 8d respectively. It was suggested that growth was delayed due to an inhibition of the germination. The effectiveness of ethanol vapour for inhibiting growth and germination of fungi is well demonstrated. However, all the experiments were carried out on products and media that contained water. Therefore, ethanol in the headspace would have been

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absorbed by these products and media. Mycelium growing at the surface would use nutrients and ethanol present in the medium. Therefore the real cause for inhibition could be the ethanol absorbed by the medium rather than ethanol vapour itself. The same phenomenon would have occurred also during germination, although to a lesser extent because interactions between the spore and the medium at the early stage of germination are limited.

4.4. Influence of ethanol on inactivation of fungal spores It was shown that at 23  C conidia of A. niger and P. notatum were completely inactivated after being exposed to 25% ethanol for 3d and 1d respectively (Bacílková, 2006). Similarly, more than 2 log inactivation of conidia of some Penicillia was achieved by applying 20% (0.7 kPa) ethanol vapour in the headspace at 25  C for 1 d. An increase of the inactivation up to 5 log was obtained for 40%, 1.5 kPa, (Dantigny et al., 2007). It was demonstrated that the effect of ethanol vapours on spore inactivation depended on temperature. An increase in temperature from 10 to 30  C was more important than an increase from 5 to 10% (w/w) ethanol for explaining the inactivation of P. digitatum and P. italicum (Dao, Bensoussan, Gervais & Dantigny, 2008). According to the ClausiuseClapeyron law, an increase in temperature caused an exponential increase in the ethanol vapour pressure. The increase in the ethanol vapour pressure can be obtained also by decreasing the water activity (Lerici et al., 1996). In fact, at reduced aw less free water is available for binding ethanol resulting in an increase of ethanol vapour pressure. Accordingly, the inactivation of spores of P. chrysogenum, P. digitatum and P. italicum was greater at 0.70 aw than at 0.90 aw (Dao et al., 2008). Therefore, the greater inactivation of these spores at reduced water activity and high temperature could be explained by an increase of the ethanol vapour pressure. The relationship between the inactivation of enzymes by ethanol and its vapour pressure was studied in a simple model containing polyphenoloxidase, PPO, (Lerici & Manzocco, 2000). Ethanol concentrations, water activity and temperature were undoubtedly the dominant factors affecting efficiency of ethanol towards PPO. It was demonstrated that the combined effects of these variables were higher than the simple addition of the single actions. This was attributed to an indirect effect of temperature (and aw) on ethanol action and supported the hypothesis that ethanol toxicity can be described by its vapour pressure. The influence of the relative humidity (70e90%) and temperature (20e30  C) at a constant ethanol vapour pressure 0.6 kPa was assessed for inactivating some Penicillia (Dantigny et al., 2007). At 70% RH, 30  C all conidia were viable after 24 h and 48 h exposure to ethanol vapours, Table 5. Overall the greatest inactivation was shown at 90% RH, 20  C. In this experimental condition the ethanol concentration in the solution was the greatest, 23.40 g/100 g solution. In contrast inactivation was not exhibited for the lowest ethanol concentration, 7.99 g/100g solution. However, there was no

365

correlation between inactivation and the concentration of ethanol in the solution. For all the species the inactivation was greater at 80% RH, 25  C (13.30 g ethanol/100g solution) than at 70% RH, 20  C (15.22 g ethanol/100 g solution). In fact a more detailed analysis has shown that RH was the key factor for explaining inactivation (Dantigny et al., 2007). Temperature was also important, but to a lesser extent. The effect of ethanol vapours (range 0.3e0.45 kPa) on inactivation of conidia obtained by a standardised protocol that consisted in preparing spore suspension was compared to that of dry-harvested conidia for some species of Penicillium. While all dry-harvested conidia remained viable after 24h treatment, about one log10, 3.5 log10 and 2.5 log10 reductions were observed for hydrated conidia of P. chrysogenum, P. digitatum and P. italicum respectively (Dao & Dantigny, 2009). These results suggested that the intracellular water activity of the conidia may be correlated to the observed differences in sensitivity to ethanol vapours. Ethanol is a very hydrophilic molecule, and more ethanol could be dissolved in a hydrated conidia during the exposure to vapours, eventually inactivating the conidia. In contrast to conidia produced at the laboratory, conidia found in the environment are usually not hydrated. It was shown that at 2.8 kPa ethanol, more than 4 log10 reductions in viable dry-harvested conidia of P. chrysogenum, P. digitatum and P. italicum were achieved after 24-h exposure (Dao, Dejardin, Bensoussan, & Dantigny, 2010).

5. Mode of action of ethanol on fungi The major target of ethanol as a stress agent is the cell membrane of fungal cells. But it has many other effects, such as denaturation of proteins (Mishra, 1993), and inhibition of the uptake of various nutrients, e.g., the non-competitive inhibition of uptake of glucose and ammonium ions (Leão & van Uden, 1982; Thomas & Rose, 1979). The inhibition of fungal growth was partly attributed to the decrease of aw due to ethanol. Water stress accounted for up to 31, 18 and 6% of growth inhibition of A. oryzae by ethanol at 25, 40, and 42.5  C respectively (Hallsworth, Nomura, & Iwahara, 1998). The main function of the cell membrane of micro-organisms is to form a permeability barrier, regulating the passage of solutes between the cell and the external environment. In the environment, micro-organisms may be confronted with lipophilic compounds such as alkanols which preferentially accumulates in membranes. This accumulation will affect physicochemical properties of membranes and consequently their functioning (Weber & de Bont, 1996). Lipophilic compounds, which possess a high affinity for the cell membrane are more toxic than less lipophilic compounds. For instance, ethanol is only toxic for micro-organisms at high concentrations (several %), whereas solvents like toluene are already toxic in the mM range (Heipieper, Weber, Sikkema, Keweloh, & de Bont, 1994). However, this correlation does not apply for hydrophobic

Table 5 Influence of relative humidity and temperature on inactivation of conidia of some Penicillium species at a constant ethanol vapour pressure Pe ¼ 0.6 kPa. Adapted from Dantigny et al. (2007). RH (%)

T ( C)

EtOH (g)

H2O (g)

Gly (g)

log (N0/Nt) mean  sd P. chrysogenum 24H

70 90 80 70 90

20 20 25 30 30

15.22 23.40 13.30 7.99 12.24

41.18 76.40 53.44 38.31 71.63

43.60 0.20 33.26 53.70 16.12

0.00 3.96 3.72 0.00 3.68

P. digitatum 48H

    

0.00 0.04 0.05 0.00 0.05

2.33 4.26 3.76 0.00 3.80

24H     

0.05 0.02 0.10 0.00 0.10

2.86 5.73 4.58 0.00 5.49

P. italicum 48H

    

0.58 0.46 0.17 0.00 0.24

2.91 6.04 5.96 0.00 6.43

24H     

0.81 0.00 0.00 0.00 0.00

2.66 4.24 4.25 0.00 4.22

48H     

0.03 0.06 0.04 0.00 0.09

2.89 4.77 4.74 0.00 5.45

    

0.19 0.08 0.10 0.00 0.04

366

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solvents that are not generally toxic for micro-organisms (Inoue & Horikoshi, 1991; Vermuë, Sikkema, Verheul, Baker, & Tramper, 1993). In fact, their poor solubility in water will prevent them from reaching high concentrations in the membrane (Osborne, Leaver, Turner, & Dunnill, 1990). The maximum concentration of a solvent in the aqueous phase decreases and the hydrophobicity increases with increasing the carbon chain length (Weber & de Bont, 1996). Depending on the hydrophobicity of the solute, it will accumulate more or less deeply into the bilayer. Alkanols like ethanol will interact with the headgroup area and alkanes with the fatty acid acyl-chains. The toxicity of alcohols is directly related to the length of their aliphatic chain and their hydrophobicity (or lipophilicity). The order of toxicity of alcohols to spores of Sclerotinia fructicola, R. stolonifer and G. persicaria was propanol, isopropanol, ethanol and methanol (Ogawa & Lyda, 1960). The addition of short-chain alcohols such as ethanol has a variety of biophysical effects. Ethanol is a small molecule, which affects the physical state and biological functions of cell membranes. It interacts with membranes at the lipidewater interface, weakening the hydrophobic barrier to the free exchange of polar molecules, thereby perturbing membrane structure and function. This could result in an increase in the surface area occupied by each phospholipids molecule and a decrease in membrane thickness, again increasing membrane permeability (Sikkema, de Bont, & Poolman, 1995). There is only one small proportion of ethanol that is distributed in the membrane. When the ethanol is intercalated within the hydrophobic part of the membrane, the polarity of this zone increases and this authorizes the passage of other polar molecules through the semi permeable membrane. Indeed, during insertion in the double-layered one on the level of the polar heads, it has been demonstrated that short-chain alkanols (C  3) can promote the formation of an unusual phospholipid aggregation structure, the interdigitated phase (Simon & McIntosh, 1984; Slater & Huang, 1988; Vierl, Löbbecke, Nagel, & Cevc, 1994). In the interdigitated phase the lipid acyl-chains from the opposing monolayers are fully interpenetrated, thereby exposing the terminal methyl groups. These short-chain alkanols will anchor with their polar moiety to the phopholipid headgroup, and with the non-polar part between the phospholipid acyl-chains, Fig. 1. Since non-polar moiety of these molecules is short compared to the fatty acid acyl-chains, these molecules would potentially cause voids between the lipid chains in the bilayer interior. As the energy of formation of holes between hydrocarbons is extremely large, the lipids respond by forming the interdigitated phase (Simon & McIntosh, 1984). As polar headgroups are important for the barrier properties of the membrane bilayer, it is expected that the formation of such lipid phase will result in an increase permeability of the membrane. Accordingly, a decrease in the osmotic water permeability of the cell membrane has been observed with increasing concentration of alcohol (25% decreases at 2% ethanol and 60% decrease at 5% ethanol) with 1.0e1.25 M ethanol required to induce loss of semipermeability (Jones, 1989). Although the interdigitated lipid phase

Fig. 1. Schematic drawing, showing the effect of ethanol on the conversion of lipid bilayers from the non-intergitated gel phase (Lb) to the fully interdigitated gel phase (LbI). Adapted from Weber and de Bont (1996).

can be induced by various additives it should be noted that this lipid aggregation structure is only observed for bilayers in the gelphase (Simon & McIntosh, 1984; Slater & Huang, 1988; Vierl et al., 1994). In micro-organisms, however, most of the phospholipids are generally in the liquid-crystalline phase. It is therefore unclear if the interdigitated phase will occur or can be induced by solvents in microorganisms. The membrane is besides a barrier also a matrix for various important enzymes. These include enzymes involved in solute transport, and enzymes participating in the electron transport chains. The results of Thomas and Rose (1979) indicate a relationship between inhibition of growth and nutrient transport by alcohol and membrane lipid composition. The alcohols themselves are highly permeable to the cell membrane, and it is proposed that the increased resistance to water flow is due to the interaction of alcohol with the membrane causing a decrease in the size of the water pores. The membrane of spore is protected by a wall made up of several layers enable him to resist adverse conditions. The wall of the spore of fungi consisted in a polymer association of polysaccharides and chitin-glucanes. The envelope of the spore contains also simple sugars (galactose, mannose), cellulose, proteins and lipids. During germination of P. chrysogenum, it was observed that at 3.5% ethanol some conidia had produced abnormal germinating tube that eventually ruptured at the apex level while other conidia were inhibited at the swelling stage (Dantigny, Tchobanov et al. 2005). These observations support the hypothesis that ethanol is responsible for the leakage of solutes across the membrane and for cell lysis following decreased peptidoglycan cross-linking in the growing cell wall (Ingram & Buttke, 1984). Relatively high concentrations of ethanol are required to kill fungal spores compared with vegetative bacteria (Larson & Morton, 1991). Unbalanced cytoplasmic permeability and cytosol leakage ultimately leading to disintegration of the cell, was reported to be efficient in ethanol concentrations varying between 50 and 80%, with a maximum at 70% (Hugo, 1971; Russell, Hugo, & Ayliffe, 1992). But mature, swollen spores (due to hydration) are more easily killed than dry and dormant ones, implying that any rational sanitizing technique aiming at sporicidal action should be whenever possible be applied to matures ones (Borick & Pepper, 1970; Tomazello & Wiendl, 1995). Denaturation caused by alcohols can affect the proteins of the cell wall, cytoplasmic membranes and proteins contained in the cytoplasm (Bacílková, 2006). According to CabecaSilva, Madeira-Lopes, and van Uden (1992), the primary sites of action of both heat and ethanol are mitochondrial membranes. Or since low concentrations of ethanol can lower the temperature at which phospholipids undergo a phase change, the increases in the spore mortality and decay control following the addition of ethanol may have resulted from a lowering of the phase-change temperature of mitochondrial membranes of the spores under these conditions (Karabulut et al., 2004). 6. Conclusions Ethanol has GRAS status in the USA, and it is an approved substance for use as a disinfectant or sanitizer in organic crop production by USDA National Organic Program (2001). Interest in developing alternatives to fungicides is high because they are being lost due to fungicide resistance, regulatory issues, and growers or consumers who may prefer not to use them (Margosan et al., 1997). In this review the effectiveness of ethanol for controlling fungi in foods was demonstrated. Many studies have shown the efficacy of ethanol treatments to reduce fruit decay. Ethanol residues after treatment of peaches and nectarines were low after storage (Mlikota Gabler et al., 2005; Margosan et al., 1997) and should pose

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a minimal regulatory issue (Anonymous, 1993). Therefore, the use of ethanol (either liquid or vapour) would have a great impact on both the economy and food quality of fruits and vegetables. The use of ethanol for preserving foods is dependent on regulations. At 2% ethanol, ethanol is not effective at controlling mould growth, but can be used as an additional barrier. It is also uncertain whether consumers would accept an ethanol emitter in the packaging. However, ethanol is used as a solvent for spraying aroma on cakes prior to packaging in the baking industry. Many fungicides are based on alcoholic molecules. Ethanol solutions at difference percentages are used as a disinfectant for personnel or surfaces. There are many indications that lower levels of ethanol were required in the vapour phase as compared to direct addition to a medium or food for complete inhibition (Lihandra, 2007; Smith et al., 1987). Ethanol is a very hydrophilic molecule. It was shown that the effect of ethanol is greatly dependent on the degree of hydration of the spore (Dantigny et al., 2007). Therefore, this factor should be carefully controlled prior to comparing the effect of ethanol solution and vapour. In cases of large volumes to be treated, or equipment susceptible to corrosion, the use of ethanol solutions is prohibited. Ethanol vapours may be an interesting alternative to ethanol solutions. One major concern with the use of ethanol vapour is its explosive potential under high vapour pressure. The lower flammability limit is 3.3 kPa (Anonymous, 1993). In this review, it was shown that 20% ethanol at 25  C (0.7 kPa) could inactivate significantly fungal spores. However, the air in manned workplaces cannot contain ethanol at more than 0.1 kPa (Anonymous, 1993). Moreover, ethanol vapour could be used for the protection of cereal grains against the degradation and the production of mycotoxin by fungi during storage. In contrast to ethanol solutions, ethanol vapour has the advantage of reaching any remote places of the silo, especially humid places where moulds were more likely to develop. This is a promising use of ethanol vapour for preventing from the development of toxigenic moulds during storage of cereals and grains. At present, fumigation with ethanol vapour for decontaminating a cooler room in a bakery was proved effective but should be optimised. These are applications that are currently being examined.

Acknowledgements Ailsa Hocking is gratefully acknowledged for helpful information on the taxonomy of Penicillium species.

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